Immunoelectron Microscopy of Parasites

Immunoelectron Microscopy of Parasites MASAMICHI AIKAWA AND CARTER T. ATKINSON

Institute of Pathology, Case Western Reserve University, Cleveland, Ohio 44106, USA

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Methods . . . . . .

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D. Immunocytochemical controls . . . . . . . . . . B. Toxoplasma, Sarcocystis and Eimeria

D. Trypanosoma and other flagellates E. Trichinella and other nematodes . . F. Schistosoma and Fasciola ................................... References

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I. INTRODUCTION lmmunocytochemistry includes the broad collection of techniques that use antibodies and suitable markers to localize specific antigens in cells or tissues ADVANCES IN PARASITOLOGY VOL 29 ISBN 0-12-031729-X

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by light or electron microscopy. Procedures which use fluorescein- or peroxidase-antibody conjugates to localize antigens by light microscopy have been widely used by parasitologists, but it is only recently that the more powerful ultrastructural techniques of immunoelectron microscopy have been applied to the study of protozoan and helminth parasites. Immunoelectron microscopy was introduced by Singer (1 959) with the development of ferritin-antibody conjugates. In the 30 years since then, refinements of the technique have made immunoelectron microscopy an indispensable tool for biological scientists. Recent applications of cell and molecular biology to parasitology have made immunoelectron microscopy increasingly important for localizing protective antigens for vaccine development and for characterizing the subcellular make-up and function of specific parasite organelles. In this review, we discuss the theory and methods of immunoelectron microscopy and summarize current applications to the study of protozoan and helminth parasites. We hope that the procedures and applications which are described in this chapter will encourage other parasitologists to use this technique for molecular and ultrastructural studies of host/parasite interactions. 11. METHODS A.

FIXATION

Fixation for immunoelectron microscopy should: (i) stabilize the antigen or antigens under investigation to prevent their diffusion or extraction from the tissue during subsequent processing; (ii) preserve the tertiary structure of the antigen or antigens to retain antigenicity and reactivity with antibodies; and (iii) preserve acceptable ultrastructure (Hayat, 1981; Bullock, 1984). In general, fixatives that preserve good ultrastructure by extensively crosslinking proteins often do so at the expense of antigenicity. The most widely used fixatives for immunoelectron microscopy are glutaraldehyde and formaldehyde. Many workers have preserved acceptable ultrastructure and antigenicity by using low concentrations of buffered glutaraldehyde (0.11 .O%) for brief time periods (10 -30 min). Higher glutaraldehyde concentrations and longer fixation times often destroy immunoreactivity, although many antigens are able to survive. Formaldehyde is superior to glutaraldehyde as a fixative for immunoelectron microscopy because its lower molecular weight allows rapid penetration of tissue and because it is able to stabilize most proteins without destroying their antigenicity (Hayat, 198 I). However, when used alone, neither low (1 YO)nor high (up to 8%) concentrations of buffered formaldehyde preserve ultrastructure well. In addition, formaldehyde fixation is, to some extent, reversible, depending on the concen-

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tration of the formaldehyde solution, length of exposure to the fixative and type of cell or tissue. Most workers have tried to combine the best qualities of glutaraldehyde and formaldehyde by using mixtures of the twegenerally 1 4 % formaldehyde and 0.05-1 .O% glutaraldehyde. These solutions give the best results when prepared fresh from paraformaldehyde powder and recently distilled monomeric glutaraldehyde. Commercial formalin solutions should be avoided because they often contain methanol and other impurities (Hayat, 1981). Since some antigens are extremely sensitive to glutaraldehyde fixation, several glutaraldehyde-free fixatives have been developed. These include carbodiimidates and bifunctional diimidoesters such as diethylmalonimidate and dimethylsuberimidate (McLean and Singer, 1970; Hassell and Hand, 1974; Yamamoto and Yasuda, 1977; Hayat, 1981) and more commonly used formaldehyde-based fixatives such as PLP (periodic acid-lysine-paraformaldehyde) fixative (McLean and Nakane, 1977) and formaldehyde-picric acid combinations (Stefanini et al., 1967). PLP fixatives were devised to preserve fine structure by the oxidation of carbohydrates with periodic acid to produce aldehyde groups which could then be cross-linked to each other or to amino groups of proteins by lysine and paraformaldehyde (McLean and Nakane, 1977). Picric acid has been used as an ingredient because it is believed to precipitate proteins without significant denaturation (Bullock, 1984). Both PLP and formaldehyde-picric acid fixatives have been modified to include glutaraldehyde (Dae et al., 1982; Newman et al., 1982; Somogyi and Takagi, 1982; Gendelman et al., 1983). The incorporation of amines such as cyclohexylamine into formaldehyde-based fixatives may also improve structural preservation without destroying antigenicity (Luther and Bloch, 1989). Antigens sensitive to glutaraldehyde may also be protected from relatively harsh conditions of fixation by the use of ethylacetimidate (Tokuyasu, 1984). This compound blocks amino groups and helps to reduce the amount of cross-linking during glutaraldehyde fixation. Secondary fixation with osmium tetroxide is not recommended for postembedding immunoelectron microscopy because it destroys the antigenicity of most cellular antigens. Rarely, the antigenicity of material fixed with glutaraldehyde-osmium and embedded in resin may be restored if gridmounted, thin sections are etched with a strong oxidizing agent such as hydrogen peroxide or sodium metaperiodate (Bendayan and Zollinger, 1983). B.

ACCESSIBILITY OF ANTIGENIC SITES

Unlike surface antigens which can be labeled by incubation of intact living or fixed cells with immunoreagents, intracellular antigens must first be made

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accessible to antibodies. A variety of pre- and post-embedding techniques has been devised to solve this problem. The choice of a particular procedure is affected by the sensitivity of the antigen to buffers, dehydrating agents and embedding media, and by the quality of structural preservation that is needed. 1. Pre-embedding immunocytochemistry

Pre-embedding immunocytochemical procedures include techniques that are applied to cells or tissues before they are embedded in a suitable supporting medium. These procedures avoid the loss of immunoreactivity that often accompanies exposure to harsh dehydration and embedding reagents, but require that intact cells and tissues be made permeable to immunoreagents. Both physical and chemical methods are available for doing this. Physical methods include: (i) infiltrating the sample with a suitable cryoprotectant, such as sucrose, glycerol or dimethylsulfoxide, and subjecting the cells to rapid freezing and thawing in liquid nitrogen; or (ii) passing the sample through an ascending and descending series of ethanol dilutions that were prepared with the rinsing or storage buffer (Somogyi and Takagi, 1982; Eldred et af., 1983). Both methods disrupt cellular membranes sufficiently to allow penetration of immunoreagents, while still preserving acceptable ultrastructure (Eldred et al., 1983; Priestley, 1984). Chemical methods include brief exposures to low concentrations of detergents such as Triton X-100 or saponin to disrupt cell membranes (Priestley, 1984). 2. Post-embedding immunocytochemistry Post-embedding techniques are used when intact cells and tissues are needed to examine the intracellular distribution of antigens. Samples are embedded and sectioned before labeling with immunoreagents. Post-embedding procedures have a number of advantages over pre-embedding techniques. Cells are not damaged while being rendered permeable, a single sample can be sectioned and tested with a variety of different antibodies, and doublelabeling techniques with particulate markers are simple to perform. (a) Cryo-ultramicrotomy. Cryo-ultramicrotomy is still the best method for localizing antigens that are sensitive to organic solvents or embedding resins. In this procedure, tissues are infused with 2 . 3 sucrose ~ and then quickly frozen in liquid nitrogen. During sectioning, ice acts as the supporting material (Griffiths er af., 1984; Tokuyasu, 1986). While this is currently the most sensitive post-embedding technique, cryo-ultramicrotomy requires specialized equipment, is technically difficult, and yields sections that have low contrast.

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(b) Resins. During the past 10 years several popular hydrophilic acrylic and methacrylate resins have been developed. These include methacrylate/ acrylate mixtures such as Lowicryl K4M, HM20, K11M and HM23 (Armbruster et al., 1982; Carlemalm et al., 1982, 1985; Roth, 1989), acrylic resins such as LR White and LR Gold (Timms, 1986; Bendayan et al., 1987; Newman and Hobot, 1987), the older methacrylates such as glycol methacrylate (Leduc and Bernhard, 1967; Cole, 1984), and mixtures of methyl methacrylate and newer acrylic resins (Escolar et al., 1988). These resins are attractive because they can be used at low temperatures where the denaturing effects of alcohols and other organic solvents are reduced (Armbruster et al., 1983). Their polar, hydrophilic nature allows them to polymerize without complete dehydration of the sample, thereby preserving hydration shells around proteins and allowing closer interaction of aqueous solutions of antibodies with the resin surface (Armbruster et al., 1982; Newman and Hobot, 1987). With glycol methacrylate and methyl methacrylateacrylic mixtures, dehydration of the tissue is possible without exposure to alcohols (Leduc and Bernhard, 1967; Escolar et al., 1988). The choice of a post-embedding procedure will ultimately depend on the sensitivity of the antigen or antigens of interest. Bendayan et al. (1987) found that embedding media had a significant effect on the localization of amylase in pancreatic tissue. Intense labeling was obtained with cryosectioned material and material embedded in glycol methacrylate, while epoxy and acrylic resins were less efficient. C.

VISUALIZATION OF ANTIGENIC SITES

Since ferritin-antibody conjugates were first introduced as electron-dense markers for immunoelectron microscopy (Singer, 1959), several different probes have been developed for the visualization of antigen-antibody reaction sites. These include particulate markers conjugated to a suitable ligand by covalent or non-covalent bonds, e.g. ferritin, colloidal gold and iron-dextran complexes, and covalently conjugated enzymatic markers that produce an electron-dense reaction product when incubated with an appropriate substrate, such as horseradish peroxidase (Horisberger, 1984). Both enzymatic and particulate marking systems have inherent advantages. Final choice of a marker will depend on the degree of resolution that is desired, on whether quantitative or qualitative information is needed, on the accessibility of the antigen to antibodies, and on the sensitivity of the antigen to fixation and embedding. 1, Enzymatic markers

Horseradish peroxidase is the most popular enzymatic marker for immuno-

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electron microscopy. Tissue-bound antibodies are visualized by exposure of the peroxidase-antibody conjugate to 3,3’-diaminobenzidene to form an osmiophilic, electron-dense reaction product (Horisberger, 1984; Sternberger, 1986). This enzyme can be coupled directly to the primary antibody and used as a one-step, direct procedure, or it can be coupled to secondary antibodies or other ligands and used as a multistep, indirect technique that significantly increases the sensitivity of the procedure. These indirect “sandwich” approaches include the peroxidase-antiperoxidase (PAP) technique and its many modifications (Ordronneau, 1982; Sternberger, 1986), the avidin-biotin-peroxidase complex (ABC) technique (Childs and Unabia, 1982; Childs, 1983a), the protein A-peroxidase technique (Dubois-Dalcq et al., 1977), the dinitrophenyl (DNP) hapten sandwich staining (DHSS) procedure (Newman and Jasani, I984), and the streptavidin-peroxidase technique (Shi et al., 1988). Immunoenzyme techniques have a number of advantages over procedures which use particulate markers. The relatively small size of most of the reagents, as compared with ferritin and colloidal gold markers, allows them to penetrate cells and some hydrophilic resins in pre- and post-embedding procedures (Zafrani et al., 1983; Newman and Hobot, 1987). In preembedding protocols, this allows the immunoreaction to be completed before dehydration and embedding steps that may destroy antigenicity. Relatively harsh fixations with osmium tetroxide may then be carried out to improve ultrastructure and contrast the reaction product. Immunoenzyme techniques are especially useful when antigen concentration is low or when only small numbers of immunoreactive sites have survived fixation and tissue processing (Pelletier and Morel, 1984). In addition, the electron-dense reaction product is often easy to spot at low magnifications. Unfortunately, immunoenzyme techniques have much lower resolution than procedures that use particulate markers. Reaction product may be artifactually deposited on structures surrounding the antigenic targets or the structures associated with specific antigens may be obscured (Courtoy et al., 1983). In addition, unstained sections must be viewed in order to distinguish clearly the reaction product from surrounding structures (Pelletier and Morel, 1984; Priestley, 1984). 2.

Particulate markers

Antibody-ferritin conjugates and colloidal gold probes which are bound to a variety of different ligands are the most popular particulate markers in current use. Iron-dextran complexes and hemocyanin have also been conjugated to immunoglobulins, but have not been widely adopted for ultrastructural studies (Miller et al., 1981; Horisberger, 1984). While the small size

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(5.54.0 nm core) and high resolution of antibody-ferritin conjugates have made them popular, their net negative charge at neutral pH may cause problems with non-specific binding to cellular structures and embedding resins in pre- and post-embedding immunocytochemical techniques (Parr, 1979; Sternberger, 1986). In addition, their small size makes them difficult to distinguish at low magnifications. The application of colloidal gold probes to electron microscopy was a major advance (Faulk and Taylor, 1971). Colloidal gold offers much greater versatility than other markers because: (i) it can be prepared in a variety of sizes for use in double- and triple-labeling experiments on the same tissue section (Slot and Geuze, 1981, 1984, 1985); (ii) it is suitable for use in both scanning and transmission electron microscopy (Hodges et al., 1987); (iii) it can be conjugated easily to a variety of ligands (Horisberger and Rosset, 1977; Hodges et al., 1984; Horisberger and Clerc, 1985; Lucocq and Baschong, 1986); and (iv) its particulate nature allows precise quantification of labeling in carefully controlled studies (Gagne and Miller, 1987; Howell et al., 1987; Kehle and Herzog, 1987; Kellenberger et al., 1987; Posthuma et al., 1987a). A variety of different ligand-gold conjugates is available for immunoelectron microscopy, including protein A-gold, avidin-gold, streptavidingold, and immunoglobulin-gold (Tolson et al., 1981; Bendayan, 1984; Roth, 1986). Protein A is a cell-wall constituent of most strains of Staphylococcus aureus that binds with high affinity to the Fc portion of immunoglobulin (Ig) (Goding, 1978; Langone, 1982). Protein A binds particularly well to IgG from hosts such as rabbits and guinea-pigs, but relatively poorly to IgG from sheep or goats or to any of the subclasses of IgG from mice (Richman et al., 1982). When mouse monoclonal antibodies are used for immunolabeling, this problem can be circumvented by conjugating the monoclonal antibody directly to colloidal gold. To avoid the preparation of separate probes for each antibody, most workers label with a secondary antibody, e.g. rabbit anti-mouse IgG, followed by incubation with protein A-gold or immunoglobulin-gold. This approach intensifies the immunolabeling and allows the use of a single protein A-gold or immunoglobulin-gold probe for a variety of different monoclonal and polyclonal antibodies. Protein G, a cell-wall protein of a human group G streptococcal strain, has been purified and used to prepare gold probes. Like protein A, it binds specifically to the Fc fragment of immunoglobulin molecules (Bendayan, 1987). Bjorck (1988) recently described protein L, an Ig-binding protein which has been isolated from the surface of Peptococcus magnus. This protein shows considerable promise for immunocytochemistry because it binds K and h light chains from all classes of immunoglobulins. Avidin, a protein isolated from egg whites, has been used for over 40 years

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as a ligand for immunocytochemical studies because of its high affinity for the coenzyme biotin. Avidin-biotin marker systems depend on the specific binding of avidin-marker conjugates to biotinylated antibodies or tissue antigens. In recent years, streptavidin has been isolated from cultures of Streptomyces avidini and used in place of avidin (Bonnard et al., 1984). Streptavidin is similar to avidin in its affinity for biotin, but is superior as a ligand because it is non-glycosylated, has a neutral PI, and is consequently less likely to bind non-specifically to tissues (Hofmann et al., 1980; Bonnard et al., 1984). Consequently, streptavidin has been used more widely than avidin in preparation of gold conjugates, although neither ligand has been as popular as protein A or immunoglobulin. D.

1.

IMMUNOCYTOCHEMICAL CONTROLS

Method specijicity

A number of non-immunological factors can cause inaccurate labeling of tissues. These factors include interaction of antibodies or probes with unquenched aldehyde groups, hydrophobic or ionic interactions of reagents with tissue components or embedding media, and pseudoperoxidase or endogenous peroxidase activity in some cells or organelles (Van Leeuwen, 1986). In addition, naturally occurring antibodies or contaminating antibodies that were raised to a poorly purified immunogen may bind to tissues to give a deceptively specific, but inaccurate, immunolocalization (Petrusz, 1983; Van Leeuwen, 1986). The following controls should be included in every labeling procedure to test the specificity of the method.

(i) Incubation with secondary antibodies and markers, or with markers alone, to establish that an immunoreaction is dependent on a complete series of sequentially added reagents. (ii) Substitution of primary antibody with preimmune serum or an irrelevant monoclonal antibody of the same class and subtype and use of a dilution series with both primary and control sera to test for tissue binding by mechanisms unrelated to antigen-antibody reactions (Buffa et al., 1979; Grube, 1980). This control will also help to determine optimal antibody concentrations and whether polyclonal sera contain naturally occurring antibodies to tissue antigens other than the one(s) under study. (iii) Whenever available, use of positive controls that were fixed and embedded by the same procedures. These help to prevent false-negative

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results and confirm that antigenic sites survived fixation and are accessible to antibodies. 2. Serum specijicity Confirming the specificity of polyclonal or monoclonal antibodies is the most important part of an immunocytochemical study. Since antibodies recognize only relatively small regions or sites on the surface of antigens, they cannot distinguish between unrelated polypeptides that have shared or cross-reacting epitopes. Polyclonal antisera may also contain contaminating antibodies which recognize antigens other than the intended one. In addition, cross-reactions are affected by: (i) the affinity of the antibodies, e.g. antibodies with low affinity for a specific antigen may be capable of binding totally dissimilar antigens; (ii) the class of the antibody, e.g. IgM antibodies have a high avidity and are “stickier” than IgG antibodies; and (iii) the structure of the antigenic target, e.g. antigens with repeating amino acid sequences seem to be more susceptible to cross reactions (Ghosh and Campbell, 1986). Another complication is the observation that the type and characteristics of the blocking agent, e.g. pH, ionic strength, detergent and protein carrier, may enhance or inhibit cross reactions (Ghosh and Campbell, 1986). Pool et al. (1983), Van Leeuwen (1982, 1986) and Van der Sluis and Boer (1986) have described controls that should be used to establish specificity of an antibody in immunocytochemical procedures. Unfortunately, a test that can unequivocally establish the specificity of a serum or antibody on intact tissues or tissue sections has not yet been devised (Van Leeuwen, 1982, 1986). Electrophoretic transfer of proteins from sodium dodecyl sulfate polyacrylamide gels to a suitable matrix, followed by an immunoblot with the serum or antibody of interest, is commonly used to establish specificity (Cumming and Burgoyne, 1985). The results may be misleading, however, since the relationships between an antigen and surrounding tissue are destroyed during extraction and purification. Since no single test can prove antibody specificity, most authors recommend using as many independent lines of evidence as possible (Childs, 1983b). These include adsorption tests with homologous or heterologous antigen to abolish immunolabeling and use of negative control tissue that lacks the antigen under investigation. When available, a panel of monoclonal antibodies to different epitopes on the same molecule may be useful for evaluating specificity; however, results may still be misleading since fixation and processing may destroy some epitopes and create other cross-reacting epitopes on unrelated molecules.

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111. APPLICATIONS TO PARASITOLOGY

A.

PLASMODIUM

1. Erythrocytic stages ( a ) Merozoite surface coat. The primary surface antigens of the Plasmodium falciparum merozoite are derived from a large precursor glycoprotein (Pf 195) that is synthesized late in schizogony and proteolytically processed at the time of merozoite release into smaller molecular weight fragments (Lyon et al., 1987; Holder et al., 1987). Several laboratories have identified fragments of 83 kDa, 42 kDa, and 19 kDa molecular mass in extracts of free merozoites that were surface-labeled by lactoperoxidase-catalyzed iodination, as well as a variety of other processing fragments that are shed from the merozoite surface (Holder and Freeman, 1984; McBride and Heidrich, 1987; Camus et al., 1987; Lyon et al., 1987). Standard transmission electron microscopy and biochemical studies have shown that most of the merozoite surface coat is shed at the time of erythrocyte invasion (Aikawa et al., 1978). Only a few processing fragments of low molecular weight are detectable in erythrocytes infected with ring forms (Holder el al., 1985a). Vaccination experiments with purified and synthetic fragments of Pf 195 have been successful in inducing partial and complete immunity to challenge infections in Aotus and Saimiri monkeys as well as humans (Cheung et al., 1986; Siddiqui et al., 1987; Patarroyo et al., 1988). Both pre- and postembedding immunolabeling with immune serum or monoclonal antibodies to Pfl95 and specific processing fragments have localized epitopes associated with this molecule at the surface of merozoites and mature schizonts (Fig. 1) (Langreth and Reese, 1979; Howard et al., 1984, 1985; Pirson and Perkins, 1985; Atkinson et al., 1987a; Chulay et al., 1987), suggesting that circulating antibodies may bind to merozoites and inhibit invasion of erythrocytes or facilitate phagocytosis of the parasites by macrophages and neutrophils. Similar high molecular weight glycoproteins have been identified in mature schizonts of P. knowlesi and P . yoelii and localized on the surface of mature schizonts and merozoites by immunoelectron microscopy (Epstein et al., 1981; David et al., 1984; Oka et al., 1984; Aikawa et al., 1986a). Aikawa et af. (1 986a) used cryo-ultramicrotomy and protein A-gold to localize the 143/140 kDa fragment of the P . knowlesi merozoite surface antigen on the endoplasmic reticulum and plasma membrane of multinucleated P. knowlesi schizonts (Fig. 2). These observations indicated that the molecule is synthesized first in the endoplasmic reticulum before being transported to

FIG. 1. LR White section of a P . falciparum merozoite (M) labeled with human hyperimmune serum and protein A-gold. Gold particles are found on the surface of the merozoite, indicating that potential protective epitopes are associated with the merozoite surface coat. x 43 000. FIG.2. Cryosection of a P . knowfesi schizont, labeled with a mouse monoclonal antibody to protective 142/140 kDa antigens. Label is associated with the surface of merozoites (M) and with the parasitophorous vacuole membrane (PVM). x 35 000. (Reproduced by permission from Aikawa et af., 1986, European Journal of Cell Biology, 41, 207-21 3.)

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the surface of developing schizonts. Interestingly, the antigen is also present on the parasitophorous vacuole membrane and Maurer’s clefts, suggesting that it is shed into the intracellular compartment during maturation of the parasite. A second family of merozoite surface antigens, distinct from Pf195 and ranging from 45 kDa to 56 kDa, has been identified by several laboratories and localized on the plasma membrane of intracellular and free merozoites by pre-embedding immunoelectron microscopy and cryo-ultramicrotomy (Stanley et al., 1985; Epping e t al., 1988; Miettinen-Baumann et al., 1988; Clark et al., 1989). In contrast to Pf195, this antigen does not appear to be processed (Clark et al., 1989). Antibodies to this molecule inhibit merozoite invasion, indicating that it may have potential value as a vaccine candidate (Miettinen-Baumann et al., 1988; Clark et al., 1989). (6) Rhoptry-microneme complex. Rhoptries and micronemes are electron-dense, membrane-bound organelles at the anterior end of malarial merozoites and sporozoites. The rhoptries consist of a pair of tear-drop shaped organelles that are connected to the apical end of the parasite by narrow ductules (Aikawa et a/., 1978). The rhoptries are believed to be attached to smaller membrane-bound, electron-dense micronemes in the apical cytoplasm to form an interconnected complex. During host-cell invasion, merozoites appear to secrete material from the rhoptry-microneme complex which perturbs the host-cell membrane/cytoskeleton and becomes incorporated into both the erythrocyte membrane and the invaginating parasitophorous vacuole membrane (Aikawa e f al., 1981a; Brown et nl., FIG. 3. LR White section of a P . falciparum merozoite, labeled with a sheep polyclonal antibody to calmodulin. Gold particles (arrow) are concentrated at the apical end (A) of the merozoite, demonstrating the presence of calmodulin. x 74 000. (Reproduced by permission from Matsumoto et al., 1987, European Journal of Cell Biology, 45, 3-3.) FIG. 4. LR White section of a P . falciparum merozoite that was treated with a calmodulin inhibitor; section labeled with sheep polyclonal antibody to calmodulin. Note absence of gold particles at the apical end (A) of the merozoite. x 56 000. (Reproduced by permission from Matsumoto et al., 1987, European Journal of Cell Biology, 45, 3-3.) FIG. 5. LR White section of an erythrocyte infected with P . falciparum, labeled with a mouse monoclonal antibody to glycophorin A. Note the absence of this erythrocyte integral membrane protein from around the parasitophorous vacuole membrane (PVM), indicating that the protein is not carried into the erythrocyte by the invading merozoite. Label is present on the erythrocyte membrane (arrows). x 26 000. (Reproduced by permission from Atkinson et al., 1987, European Journal of Cell Biology, 45, 192-199.)

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1985; Mikkelsen et al., 1988; Sam-Yellowe et al., 1988). Localization of calmodulin in the rhoptries by immunoelectron microscopy suggests that the process of attachment and invasion of the host erythrocyte is dependent on calcium ions (Figs 3 and 4) (Matsumoto et al., 1987). Interestingly, the parasitophorous vacuole membrane that surrounds intraerythrocytic parasites is devoid of erythrocyte cytoskeletal and integral membrane proteins,

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suggesting that it either originates from rhoptry-microneme secretions or develops from an extensively modified erythrocyte plasma membrane (Fig. 5) (Aikawa et al., 1978, 1981a; Atkinson et al., 1987b; Dluzewski et al., 1988). In 1975, Bannister et al. described microspheres (also called dense granules) at the apical end of P . knowlesi merozoites. These organelles are round, measure 50-80 nm in diameter and appear to be intermediate in size between the rhoptry and microneme. During late stages of erythrocyte invasion, microspheres move to the periphery of merozoites, fuse with the pellicle and release their contents into the parasitophorous vacuolar space. This causes further invaginations of the parasitophorous vacuole membrane (Bannister et al., 1975). This process appears to be identical to the fusion of dense granules with the pellicle of Sarcocystis sporozoites soon after host-cell invasion and the subsequent release of electron-dense material into the parasitophorous vacuole space (Entzeroth et al., 1986). Recently Torii et al. 1989b performed immunoelectron microscopy with antibodies which react specifically with P . knowlesi microspheres. In extracellular merozoites, gold particles were associated only with microspheres (Fig. 7(a)), while no labeling was observed on rhoptries or micronemes. During erythrocyte invasion, microspheres moved to the surface of merozoites. Gold label was aggregated at the outside of the merozoites at the point where microspheres released the contents (Fig. 7(b)) into the parasitophorous vacuole space. The parasitophorous vacuole membrane adjacent to these areas formed elongated finger-like channels into the host-cell cytoplasm. Gold particles extended from the surface of merozoites into these channels, indicating that contents of the microspheres may have caused their formation. Although discharge of microsphere contents has not been described, these organelles also appear to be present in the merozoites of other species of Plasmodium. Since micronemes and microspheres are difficult to FIG. 6. LR White section of a P . fakiparum merozoite labeled with immune serum from an Aotus monkey which had been vaccinated with a 102/132/143 kDa rhoptry complex. Gold particles are associated with the rhoptries (R), confirming the subcellular location of this immune target. x 46 500. (Reproduced by permission from Atkinson et al., 1987a, Journal of Parasitology 73, 1235-1240.) FIG. 7. (a) LR White section of a P. knowlesi merozoite labeled with antibodies specific to microspheres (M). Label is associated only with microspheres (M), not with rhoptries (R)and micronemes (Mi). ~ 3 4 0 0 0 .(b) LR White section of P . knowlesi just after host cell invasion, labeled similarly. A microsphere is attached to the merozoite membrane (arrow) and the content of the microsphere is discharged to the parasitophorous vacuole space (PV), resulting in an aggregation of gold particles outside the merozoite (arrow head). Other microspheres (M) inside the parasite are also labeled with gold particles. x 55 000.

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distinguish morphologically, antigens which were thought to be present in micronemes in the past may actually be present in microspheres. Monoclonal and polyclonal antibodies and immunoelectron microscopy have been used to characterize both low and high molecular mass molecules in the rhoptry-microneme complex of P. falciparum merozoites. These consist of 240/225 kDa molecules (Roger et al., 1988), a 155 kDa molecule known as the ring-infected erythrocyte surface antigen (Pfl55/ RESA) that has been localized specifically in micronemes (Brown et al., 1985), 130/150kDa molecules that share common epitopes with Pf155/ RESA (Uni et al., 1987), a family of 105-145 kDa molecules (Fig. 6) (Holder et al., 1985b; Atkinson et al., 1987a; Sam-Yellowe et al., 1988; Cooper et al., 1988), and a group of molecules of lower mass, 40-80 kDa (Bushel1 et al., 1988; Ingram et al., 1988; Smythe et al., 1988). Antibodies which immunoprecipitate 80/66/42 kDa and 140/130/ 105 kDa rhoptry molecules can inhibit merozoite invasion in vitro (Schofield et al., 1986; Cooper et al., 1988). Both RESA and a complex of 102/132/143 kDa rhoptry proteins are partially protective when used to vaccinate Aotus monkeys (Collins et al., 1986; Siddiqui et al., 1987). The potential importance of these antigens in stimulating protective immunity is supported by the observation that human hyperimmune serum to P.falciparum contains antibodies which bind specifically to the rhoptry-microneme complex. Rhoptry-microneme antigens have also been localized in merozoites of P. yoelii and P . brasilianum. Oka er al. (1984) localized a 235 kDa antigen in the rhoptries of P. yoelii merozoites which is capable of inducing protective immunity in mice. Torii et al. (1989a) used immunoelectron microscopy and a panel of three monoclonal antibodies to localize 18/16/14 kDa microneme antigens of P. brasilianum in merozoites and ring-infected erythrocytes. Immunolabeling was detected in the micronemes of extracellular merozoites, on the erythrocyte membrane of recently invaded red blood cells, and on knobs and caveolae that developed in the infected red blood cell membrane as the parasites matured. These observations suggest that some microneme antigens are inserted into the plasma membrane of the infected erythrocyte at the time of host-cell invasion, where they may be modified during parasite development into knobs and caveolae. Studies of Pf 155/RESA of P. falciparum merozoites have shown that this molecule is translocated from the micronemes of invading merozoites to the erythrocyte membrane/cytoskeleton (Fig. 8) (Brown et al., 1985). Uni et al. (1987) localized a related 130/150 kDa molecule of P . falciparum in micronemes of merozoites and on the membrane of newly-invaded erythrocytes, suggesting that it is secreted into the membrane by merozoites at the time of invasion. Unlike the situation with P. brasilianum, the antigens were no longer detectable on the erythrocyte membrane after the parasites

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FIG.8. LR White section of an erythrocyte infected with a P.falciparum ring stage parasite (P). The section was labeled with a rabbit antiserum to a synthetic repeat region of Pf155/RESA. Gold label on the infected erythrocyte membrane (arrows) indicates the presence of Pf155/RESA. x 13 000. Inset: higher magnification showing Pf I55/RESA on the infected erythrocyte membrane. x 42 000.

developed into trophozoites. Uni et al. (1987) found that gold particles became more numerous within the parasite and the erythrocyte cytoplasm adjacent to the parasite. Significant labeling was found in the erythrocyte cytoplasm adjacent to mature gametocytes. A similar study using immunoelectron microscopy and polyclonal and monoclonal antibodies to Pf 155/ RESA found dense immunolabeling around mature and differentiating gametocytes of P . falciparum, suggesting that this molecule plays an important role in gametogenesis (Figs 9-11) (Quakyi er al., 1989). Within five minutes after the induction of gametogenesis, clear spaces surrounded by Pf 155/RESA formed in the erythrocyte cytoplasm around developing gametes (Fig. 10). These spaces extended from a region immediately adjacent to the parasite to the erythrocyte membrane. Fifteen minutes after the induction of gametogenesis, gametes were extracellular or within a lysed erythrocyte membrane that was densely labeled by antibody to Pf 155/RESA and colloidal gold. Labeling of lysed erythrocytes was associated with the

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former stroma of the erythrocyte and with the erythrocyte membrane and the parasite plasmalemma (Fig. 11). This study indicates that Pf 155/RESA may have important functions associated with the release of gametes from host cells.

FIG.9. LR White section of an erythrocyte infected with a P . falciparum gametocyte (G), labeled with a human monoclonal antibody which identifies Pf155/RESA. Note the presence of Pfl55/RESA (arrows) in the erythrocyte cytoplasm adjacent to the parasite. x 8250. (Reproduced by permission of the American Society for Microbiology from Quakyi et al., 1989, Infection and Immunity, 57, 833-839.) Inset: higher magnification showing Pf155/RESA in the erythrocyte cytoplasm. x 38 000. FIG. 10. LR White section of a P . falciparum gametocyte (G) fixed within 5min after induction of gametogenesis. Pfl55/RESA (arrows) is associated with clear spaces (S) within the erythrocyte cytoplasm. x 21 000. (Reproduced by permission of the American Society for Microbiology from Quakyi et al., 1989, Infection and Immunity, 57, 833-839.) FIG. I 1. As gametogenesis progresses, gold particles associated with Pf 155/RESA become scattered throughout the lysed erythrocyte in association with remnants of the erythrocyte cytoplasm (EC) and the erythrocyte membrane (EM), suggesting that this protein is important in release of gametes from host erythrocytes. G, P . falciparum gametocyte. x 29 000. (Reproduced by permission of the American Society for Microbiology from Quakyi et al., 1989, Infection and Immunity, 57, 833839.) x 29 000.

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(c) Exoantigens. Exoantigens are a diverse group of soluble malarial proteins found in culture supernatants and in the plasma of infected hosts. Their relationships to one another are still poorly understood. Exoantigens that have been localized by immunoelectron microscopy are found in the parasitophorous vacuole space, Maurer’s clefts and erythrocyte cytoplasm around schizonts and mature segmenters. They appear to be released into

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culture supernatants or plasma when infected erythrocytes rupture and release merozoites, although at least one 72 kDa histidine-rich exoantigen (PfHRP2) is secreted by intact cells (Howard et al., 1986). S-antigens are the largest group of exoantigens which have been characterized, and consist of a family of serologically diverse, heat-stable proteins of large but variable molecular weights and poor immunogenicity (Wilson et al., 1975; Anders et al., 1983; Cowman et al. 1985). Immunoelectron microscopy has localized these proteins in the parasitophorous vacuole space around maturing schizonts and in Maurer’s clefts (Ingram et al., 1988; Mattei et al., 1988). Perkins (1984) identified a soluble, heat-stable molecule with mass = 130 kDa in culture supernatants of P . falciparum which bound glycophorin and which could be localized on the surface of merozoites by immunoelectron microscopy. A repeat sequence of 50 amino acids on this molecule formed a binding domain for glycophorin (Kochan et al., 1986). Antibodies to this molecule inhibited merozoite invasion, suggesting that it might be involved in the initial recognition and binding of merozoites to uninfected erythrocytes. More recently, Bianco et al. (1987) and Bonnefoy et al. (1988) described complementary deoxyribonucleic acid (cDNA) clones which encode portions of this molecule and localized it by post-embedding immunoelectron microscopy within the erythrocyte cytoplasm of schizontinfected cells. Only a minor proportion of the protein appeared to bind to the surface of intracellular and extracellular merozoites. Bonnefoy et al. (1988) suggested that this protein might be a distinct type of S-antigen which was much less abundant than polymorphic S-antigens. Its release into the blood plasma at the time of schizont rupture might bind antibodies that would otherwise inhibit merozoite invasion. Delplace et al. (1987) detected a 50 kDa antigen in supernatants from cultures of P . fakiparum. Monoclonal antibodies to this antigen identified a 126 kDa polypeptide (P126) which was processed at the time of merozoite release to 50 kDa, 47 kDa, and 18 kDa polypeptides. Knapp et al. (1989) used antibodies to a protective 140 kDa antigen of P . falciparum to isolate a gene which codes an analogous 113 kDa polypeptide with a characteristic stretch of serine residues (SERP I). Both laboratories were able to localize this antigen in the parasitophorous vacuole space and Maurer’s clefts of schizont-infected erythrocytes, suggesting that it plays some role in the release of merozoites. Cochrane et al. (1988) identified a 120 kDa antigen of P . brasilianum which has an intracellular distribution similar to P126. This antigen first appears as discrete aggregates within the host cell cytoplasm around late trophozoites. In immature schizonts and segmenters, abundant labeling becomes associated with the parasitophorous vacuole space around budding merozoites (Fig. 12). Labeling in mature segmenters spreads from

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the parasitophorous vacuole space into the erythrocyte cytoplasm, suggesting that this antigen, like P126, may have proteolytic functions that are important in the final stages of merozoite budding and the disruption and breakdown of the erythrocyte membrane.

FIG. 12. LR White section of a P . brusifiunum schizont, labeled with a mouse monoclonal antibody to a 120 kDa antigen. Gold particles are associated with the parasitophorous vacuole space (PVS) and erythrocyte cytoplasm around the mature merozoites (M). x 27 000. (Reproduced by permission of the American Society for Microbiology from Cochrane et uf., 1988, Infection and Immunity, 56, 208C2088.)

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( d ) Host-cell alterations induced by Plasmodium. The erythrocytic stages of malarial parasites induce significant morphological and functional changes in infected erythrocytes including structural alterations, i.e. knobs, clefts, caveolae, and caveolae-vesicle complexes, and changes in the composition and antigenic characteristics of the erythrocyte membrane/cytoskeleton (Sherman, 1985). Morphological changes have been recognized by light microscopy for many years and have been called by various names such as Schuffner’s dots, Maurer’s clefts, Ziemann’s stippling and Sinton’s and M ulligan’s stippling. Immunoelectron microscopy has recently demonstrated that these changes are related to the trafficking of malarial proteins from the parasite to the erythrocyte membrane/cytoskeleton (Aikawa, 1988a). Some of these alterations in host-cell morphology appear to be involved in the development of malaria-related complications in the host (e.g. cerebral malaria). (i) Plasmodium falciparum. Several changes in host cell morphology are induced by infection with P. falciparum. Knob-like protrusions develop in the erythrocyte membrane/cytoskeleton and clefts with associated electrondense material appear in the erythrocyte cytoplasm as parasites mature. Knobs are conical 40 nm protrusions of the erythrocyte plasma membrane which are underlain by electron-dense material. Knobs possess cytoadherence activity and often form focal junctions with the endothelial cells and adjacent erythrocytes (Aikawa, 1988b). At least four malarial proteins (HRPl, HRP2, EMPl and EMP2) have been identified on the surface or in association with the cytoskeleton of erythrocytes infected with P.falciparum. HRPl is a 90 kDa histidine-rich protein that has been localized by immunoelectron microscopy within electron-dense knob material and in the clefts (Ardeshir et al., 1987; Culvenor et al., 1987; Pologe et al., 1987; Taylor et al., 1987). It appears to be related to the structural formation of the knob (Ardeshir et al., 1987). HRP2 is a different, 70 kDa water-soluble histidinerich protein which has been localized by immunoelectron microscopy within the erythrocyte cytoplasm and in association with Maurer’s clefts and the erythrocyte plasma membrane. In contrast to HRPl, which is tightly bound to the erythrocyte cytoskeleton, HRP2 is secreted by infected cells (Howard et al., 1986). EMPl and EMP2 d a not contain histidine and both are proteins of 300 kDa (Leech et al., 1984; Howard et al., 1987). Although EMPl appears to have cytoadherence activity (Leech et al., 1984), immunoelectron microscopy has so far failed to demonstrate its presence on the surface of the knobs. EMP2 has been localized in the parasitophorous vacuole of schizonts, within membrane-bound vesicles in the erythrocyte cytoplasm and in association with knobs and the cytoplasmic side of the erythrocyte membrane/cytoskeleton (Coppel et al., 1986; Howard et al., 1987).

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Several recent studies have shown that both malarial and host-cell antigens are associated with the Maurer’s clefts of erythrocytes infected with P . falciparum. Atkinson et al. (1987b) studied the distribution of ankyrin (Fig. 13), spectrin, band 3 (Fig. 14), and glycophorin A (Fig. 5 ) in erythrocytes infected with P . falciparum, to determine whether movement of parasite proteins and membrane vesicles between the parasitophorous vacuole membrane and erythrocyte surface membrane involves internalization of erythrocyte membrane/cytoskeletal proteins. No changes were detected in the distribution of spectrin, band 3 or glycophorin A in erythrocytes infected with ring forms, trophozoites, or schizonts. However, ankyrin was localized on the cytoplasmic face of flattened Maurer’s clefts in erythrocytes infected with trophozoites or schizonts (Fig. 13). Ankyrin was

FIG. 13. LR White section of an erythrocyte infected with P . falciparum, labeled with a rabbit antiserum to ankyrin. Label (arrow) is associated with Maurer’s clefts (C) in the erythrocyte cytoplasm, indicating that this erythrocyte cytoskeletal protein is redistributed to the clefts during development of the parasite. x 45 500. (Reproduced by permission from Atkinson et al., 1987, European Journal of Cell Biology, 45, 192-199.) FIG. 14. LR White section of an erythrocyte infected with P . falciparum, labeled with a rabbit antiserum to band 3. In contrast to ankyrin, the distribution of this erythrocyte integral membrane protein does not change within infected red blood cells. Label (arrow) is associated with the erythrocyte membrane. No label is seen in association with clefts (C). x 38 000. (Reproduced by permission from Atkinson et d.,1987, European Journal of Cell Biology, 45, 192-199.)

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not detected on a second population of circular or vesicular clefts that appeared to originate directly from the parasitophorous vacuole membrane. The presence of ankyrin on short, flattened Maurer’s clefts supports the idea that the clefts connect to the surface of erythrocyte and may function in the transport of knob-associated electron-dense material. Malarial proteins that have been localized on Maurer’s clefts in erythrocytes infected with P. falciparum include exoantigens. e.g P126 and HRP2 (Howard et al., 1986; Delplace et al., 1987; Ingram et al., 1988), knobassociated HRPl (Culvenor et al., 1987), EMP2 (= MESA) (Coppel et al., 1986; Howard et al., 1987), and 15/19 kDa and 46 kDa molecules which appear to be integral parts of the cleft membranes (Hui and Siddiqui, 1988; Kara et al., 1988a,b). Both Hui and Siddiqui (1988) and Kara et al. (1988b) suggested that these molecules may play a role in transport of parasite antigens to the erythrocyte membrane/cytoskeleton. (ii) Plasmodium brasilianum. This species is a quartan malarial parasite of new world monkeys which has morphological, immunological and genetic similarities to P. malariae of man (Sterling et al., 1972; Atkinson et al., 1987c; La1 et al., 1988). Many monoclonal antibodies that have been produced to P. brasilianum cross-react with P . malariae, supporting the idea that these two organisms are closely related (Cochrane et al., 1988; Nagasawa et al., 1988). P. brasilianum, like P . falciparum, causes the formation of knobs and cytoplasmic clefts within infected erythrocytes. Cytoplasmic clefts are membrane-bound and can be divided into three types, namely short clefts, long FIG. 15. LR White section of an erythrocyte infected with P . brasilianum, labeled with a mouse monoclonal antibody to a 38 kDa antigen. Label is on short clefts (SC), but not on long clefts (LC), indicating an antigenic difference between these two populations of membranous structures. x 31 000. (Reproduced by permission of the American Society for Microbiology from Cochrane et al., 1988, Infection and Immunity, 56, 2080-2088.) FIG. 16. LR White section of an erythrocyte infected with P . brasilianum, labeled with a mouse monoclonal antibody to a 16 kDa antigen. The distribution of this molecule is restricted to long clefts (C) and the parasitophorous vacuole membrane (PVM), suggesting a close relationship. x 22 000. (Reproduced by permission of the American Society for Microbiology from Cochrane et al., 1988, Infection and Immunity, 56, 208k2088.) FIG. 17. LR White section of an erythrocyte infected with P . brasilianum, labeled with a mouse monoclonal antibody to 18/16/14 kDa antigens. Label is associated with electron-dense knobs (K) in the erythrocyte membrane. x 37 000. (Reproduced by permission of the American Society for Microbiology from Cochrane et al., 1988, Infection and Immunity, 56, 2080-2088.)

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clefts, and circular clefts. Short clefts are 0.34.8 pm long slit-like structures that are scattered randomly in the cytoplasm of infected erythrocytes. Long clefts are slightly curved or undulating and are 3-5 pm long. Some long clefts are connected with the parasitophorous vacuole membrane. Circular clefts form oblong loops in the erythrocyte cytoplasm and have a long axis of 1.5-2.3 pm. The morphological relationships among these types of clefts is unclear, but it has generally been assumed that they develop as a single interconnected network from the parasitophorous vacuole membrane. Recent work has shown, however, that they differ in antigenic composition (Cochrane et al., 1988). Monoclonal antibodies which recognize a 38 kDa antigen of P . brasilianum label short clefts (Fig. 15) but not long or circular clefts. By contrast, monoclonal antibodies which recognize a 16 kDa antigen label both long and circular clefts and the parasitophorous vacuole membrane and space (Fig. 16), indicating that these structures have a similar antigenic composition (Cochrane et al., 1988). The fact that the long-cleft antigen is associated with the parasitophorous vacuole, whereas the short-cleft antigen is not, suggests that there are regional differences in the distribution of cleft antigens or two or more distinct populations of cytoplasmic clefts. Threedimensional maps of the intracellular distribution of these antigens may eventually help to answer this question. It is clear, however, that structural and functional relationships between these populations of clefts may be more complicated than previously recognized. Maurer’s clefts are believed to function in the transport of electron-dense knob material erythrocytes infected with P . falciparum (Aikawa et al., 1986b). It is unclear, however, what role cytoplasmic clefts play in the transport of knob material in erythrocytes infected with P . brasilianum. As discussed earlier, Torii et al. (1989a) found that 18/16/ 14 kDa microneme antigens of P . brasilianum appear to be inserted into the erythrocyte membrane during merozoite invasion. These same antigens have been localized in knobs of erythrocytes infected with P . brasilianum, but do not appear to be present on clefts (Figs 15-17). It is possible that the nature of FIG. 18. LR White sections of an erythrocyte infected with P . vivax, labeled with a mouse monoclonal antibody to an 86 kDa antigen, which is associated with vesicles of the caveola-vesicle complexes (arrows). x 48 000. (Reproduced by permission from Udagama et al., 1988, American Journal ofPathology, 131,48-52.) Inset: higher magnification of a caveola-vesicle complex. Gold label is associated with vesicles (V), but not caveola (C). x 60 000.

FIG. 19. LR White section of an erythrocyte infected with P . vivax, labeled with a mouse monoclonal antibody to a 28 kDa antigen which is associated with clefts (C) and caveola-vesicle complexes (CV), suggesting movement between the two. x 63 000.

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antigens being transported to the erythrocyte surface and the antigenic composition of the clefts may vary with the stage of parasite development.

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(iii) Plasmodium vivax. Erythrocytes infected with P . vivax exhibit a fine stippling composed of reddish dots when stained with Romanovsky stains and examined by light microscopy. Electron microscopy has shown that these dots correspond to caveolae-vesicle (CV) complexes in the membrane/ cytoskeleton of infected erythrocytes. CV complexes are composed of alveolar-like clusters of small vesicles that are connected to the base of invaginations or caveolae in the erythrocyte membrane (Aikawa et al., 1977). P . vivax also causes the development of clefts within the cytoplasm of infected erythrocytes. Recently, investigators have produced a series of monoclonal antibodies against antigens of P . vivax erythrocytic stages (Udagama et al., 1988; Matsumoto et al., 1988a). Monoclonal antibodies which identify 95kDa and 86kDa P . vivax proteins produce a stippled pattern by immunofluorescence microscopy similar to the Schuffner’s dots seen in Romanovsky-stained erythrocytes infected with P . vivax (Udagama et al., 1988; Matsumoto et al., 1988a). Other monoclonal antibodies which react with a 28kDa protein produce a linear pattern when viewed by immunofluorescence microscopy (Matsumoto et al., 1988a). Immunoelectron microscopy has been used to identify the precise location of these P . vivax antigens. The 95 kDa and 86 kDa proteins are associated with vesicles of the CV complex and with scattered vesicles in the erythrocyte cytoplasm (Fig. 18) (Udagama et al., 1988; Matsumoto et al., 1988a). By contrast, the 28 kDa protein is associated primarily with cytoplasmic clefts, but is also found in vesicles of the CV complex (Fig. 19) (Matsumoto et al., 1988a). A double-labeling technique clearly demonstrated that both 28 and 95 kDa antigens occur in vesicles of the CV complex, suggesting that the vesicles bud from cytoplasmic clefts and then move through the erythrocyte cytoplasm to the host cell surface. Thus, both clefts and CV complexes appear to be involved in the trafficking of P . vivax antigens from the parasite to the erythrocyte membrane. 2. Sporogonic stages

Immunization with irradiated sporozoites produces a considerable degree of protection against rodent, simian and human malaria. This protection is mediated in part by antibodies which neutralize sporozoites, abolish their infectivity, and cause the formation of a thread-like precipitate on the surface of parasites. This reaction has been named the circumsporozoite precipitation (CSP) reaction (Vanderberg et al., 1969). Ultrastructural examination of sporozoites that have been incubated with mouse immune serum and an anti-mouse immunoglobulin conjugated with hemocyanin demonstrated the importance of antisporozoite antibodies in the formation of the CSP reaction (Figs 20, 21) (Cochrane et al., 1976).

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FIG.20. Epon section of a sporozoite (S) that was incubated in hyperimmune serum and then fixed and processed for electron microscopy. Antibodies have bound to the surface of the sporozoites, creating a precipitate (arrows) which is sloughed by living parasites. This has been termed the circumsporozoite reaction. x 20 000 (Reproduced by permission from Cochrane et al., 1976, Journal of Immunology, 116, 859-867.)

FIG. 21. Epon section of a sporozoite (S) that was incubated in hemocyaninlabeled antisporozoite antibodies and then fixed and processed for electron microscopy. Presence of angular hemocyanin crystals (arrows) on the surface of the sporozoite demonstrates the role of antibodies in the formation of the circumsporozoite precipitate reaction. x 16 000.

Since the study by Yoshida et al. (1981), monoclonal and polyclonal antibodies to sporozoite antigens from several species of rodent, simian, and human Plasmodium have been produced. Most of these antibodies recognize a polypeptide known as the circumsporozoite (CS) protein which completely coats the surface of sporozoites. CS proteins contain an immunodominant B-cell epitope composed of tandemly repeated sequences of amino acids

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(Nussenzweig and Nussenzweig, 1985). Numerous studies have shown that antibodies to this polypeptide can neutralize sporozoite infectivity. Because of the importance of CS protein as a vaccine candidate, several workers have used immunoelectron microscopy to trace its origins and expression throughout the life cycle of malarial parasites. Immunoelectron microscopy of the sporogonic development of P . falciparum, P . ovule, P . mulariae and P . berghei with monoclonal antibodies to the CS proteins of each species has shown that this antigen is abundant in developing oocysts ‘and appears early in sporogony before the differentiation of sporozoites (Hamilton et al., 1988a; Nagasawa et al., 1987, 1988; Posthuma et al., 1988). Specific labeling first appeared on the plasma membrane and cytoplasm of 5to 6-day old oocysts before peripheral vacuolization of the oocyst cytoplasm and formation of sporoblasts. Labeling became denser on the plasma membrane and inner surface of peripheral vacuoles and clefts as the oocyst cytoplasm was subdivided into sporoblasts. At the same time, labeling could be detected on perinuclear membranes and endoplasmic reticulum in the oocyst cytoplasm, indicating that the CS proteins are synthesized in the cytoplasm of sporoblasts before export to the plasma membrane (Fig. 22). As developing sporoblasts contracted away from the oocyst capsule, significant labeling remained associated with its inner surface as well as with material in the subcapsular space, indicating that CS antigen sloughs from the plasma membrane of developing sporoblasts. Similar sloughing of CS antigen has been observed from mature salivary gland sporozoites (Posthuma et al, 1987b), and may play an important role in mediating the gliding motility of sporozoites (Stewart and Vanderberg, 1988). As sporozoites bud from the surface of sporoblasts, they are uniformly covered with CS antigen on the outer surface of their plasma membranes (Fig. 23). Studies of P . berghei have shown that all three membranes of the sporozoite pellicle contain CS protein (Aikawa et al., 1981b; Hamilton et al., 1988a; Atkinson et al., 1989a). Mature oocyst sporozoites as well as salivary-gland sporozoites are labeled on their surface with equal intensity by antibodies to the CS protein and also exhibit internal labeling on micronemes, perinuclear FIG. 22. LR Gold section of a P. ovule oocyst labeled with a mouse monoclonal antibody to the P. ovule circumsporozoite protein, which is located on endoplasmic reticulum (ER) and the plasma membrane (PM) of a developing sporoblast. x 22 000. (Reproduced by permission of the American Society for Microbiology from Nagasawa et al., 1987, Infection and Immunity, 55, 2928-2932.) FIG.23. LR Gold section of a P. ovule oocyst labeled with a mouse monoclonal antibody to the P. ovule circumsporozoite protein. Gold label (arrows) is associated with the surface of the sporozoites (S). x 42 000. (Reproduced by permission of the American Society for Microbiology from Nagasawa et a[., 1987, Infection and Immunity, 55, 2928-2932.)

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membranes and Golgi apparatus (Fine et al., 1984; Hamilton et al., 1988a; Nagasawa et al., 1988; Posthuma et al., 1988). These findings suggest that continuous synthesis of CS protein by sporozoites may enhance their infectivity. In contrast to the merozoite stages of Plasmodium, little is known about

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the antigenic composition of the the rhoptry-microneme complex of sporozoites. Immunoelectron microscopy of P . knowlesi, P . malariae and P . herghei sporozoites with monoclonal antibodies to the CS protein and colloidal gold has localized this antigen within micronemes (Fine et al., 1984; Nagasawa et al., 1988; Atkinson et al., 1989a). Fine et al. (1984) found labeling on the membranes of micronemes in P . knowlesi sporozoites and occasionally on rhoptries, and speculated that the antigen may be processed and stored within these organelles before movement to the sporozoite surface. The recent finding that the CS protein is continuously secreted and sloughed by extracellular sporozoites (Stewart and Vanderberg, 1988) supports the idea that the micronemes may serve as an intracellular reservoir for this molecule. 3. Exoerythrocytic stages It is only recently that improvements in culture techniques in vitro have made the exoerythrocytic stages of malarial parasites available for antigenic analysis. Most studies have used immunofluorescence microscopy and antibodies to antigens from erythrocytic parasites to trace their expression in exoerythrocytic schizonts (Hollingdale et al., 1983; Aley et al., 1987a; Szarfman et al., 1988). Studies by immunoelectron microscopy have focused on this protein. Unlike the merozoite surface coat, which is mostly shed when merozoites invade erythrocytes, all or most of the CS protein of malarial sporozoites is carried into host hepatocytes. Aley et al. (1987b) used sera against four FIG.24. LR Gold section of a P. falciparum sporozoite ( S ) within a recently invaded HepG2-A 16 hepatoma cell (H), labeled with a rabbit antiserum to the repeat region of the P. falciparum circumsporozoite protein. Gold label is associated with the surface of the sporozoite, but not with the surrounding parasitophorous vacuole membrane (PVM). ~ 4 2 0 0 0 .(Reproduced by permission from Aley et a!., 1987, Journal of Parasitology, 73, 1241- 1245). FIG. 25. LR Gold section of a P. vivax sporozoite ( S ) within a recently invaded HepG2-AI6 hepatoma cell, labeled with a mouse monoclonal antibody to the P. vivax circumsporozoite protein. In contrast to P. falciparum, label is associated with both the sporozoite surface (arrow) and the surrounding parasitophorous vacuole membrane (PVM). x 42 000. FIG.26. LR Gold section of a P. berghei sporozoite ( S ) within a recently invaded HepG2-AI6 hepatoma cell, labeled with a mouse monoclonal antibody to the P. berghei circumsporozoite protein. Gold particles are on the sporozoite pellicle (SP) as well as on the surrounding parasitophorous vacuole membrane (PVM) and space. Micronemes (M) are also labeled with gold particles. x 72 000. (Reproduced by permission of the American Society of Tropical Medicine and Hygiene from Atkinson, 1989, American Journal of Tropical Medicine and Hygiene, 41, 9-17.)

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defined regions of the P . falciparum CS protein, including both repeat and non-repeat regions, to determine whether or not particular regions of the CS protein behave differently during the invasion process. Antisera to each of the four regions bound specifically to the outer surface and pellicle of sporozoites after invasion of HepG2-A16 hepatoma cells, but not to the plasma membrane or surface of infected host cells or to the parasitophorous vacuole membrane around the intracellular sporozoites (Fig. 24). Studies of hepatoma cell invasion by P . vivax and P . berghei sporozoites using antibodies to the repeat region of their respective CS proteins have shown

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that CS protein is carried into host cells by these species also (Atkinson et al., 1989a). In contrast to P. falciparum, however, CS protein could also be localized on the parasitophorous vacuole membrane (Figs 25 and 26). Interestingly, P. falciparum sporozoites are not able to grow in HepGZA16 hepatoma cells, while both P. vivax and P . berghei can complete exoerythrocytic schizogony in these cells. This suggests that interaction of CS protein with the parasitophorous vacuole membrane may be important in mediating early stages of intracellular development. After entry of sporozoites, CS protein has been detected on the surface of developing exoerythrocytic schizonts in cultures of P. falciparum, P. vivax, P . cynomolgi, and P . berghei (Figs 27 and 28) (Aley et al., 1987a; Hamilton et al., 1988b; Suhrbier et al., 1988; Szarfman et al., 1988; Atkinson et al., 1989a,b). CS protein persists throughout exoerythrocytic schizogony in cultures of P. fakiparum, P. vivax, and P . berghei (Fig. 28) and can be detected on the surface of exoerythrocytic merozoites by immunofluorescence microscopy. By contrast CS antigen appears to disappear during exoerythrocytic development of P. cynomolgi (Atkinson et al., I989b). Immunoelectron microscopy of P. berghei has confirmed the presence of CS protein on .the surface of exoerythrocytic merozoites (Suhrbier et al., 1988; Atkinson et al., 1989a). CS protein has not been found on erythrocytic merozoites, indicating that these two populations of merozoites differ in antigenicity. Nussenzweig and Nussenzweig (1985) speculated that persistence of CS antigen during exoerythrocytic development may help maintain a state of premunition, thus allowing infected hosts to escape overwhelming infections in hyperendemic areas where exposure to sporozoites is high. Studies of P. berghei by immunoelectron microscopy have shown that CS protein is associated with the parasitophorous vacuole membrane and space and with the schizont plasma membrane during early stages of development FIG. 27. LR Gold section of a P . cynomolgi exoerythrocytic form within a rhesus monkey hepatocyte, labeled with a mouse monoclonal antibody to the P . cynomolgi circumsporozoite protein. Gold label (arrows) is associated with the surface of the parasite and the surrounding parasitophorous vacuole membrane. N, nucleus. x I8 000. (Reproduced by permission of the American Society of Tropical Medicine and Hygiene from Atkinson et al., 1989, American Journal of Tropical Medicine and Hygiene, 40, 131-140.)

FIG.28. LR Gold section of a P . berghei exoerythrocytic schizont within a HepG2A16 hepatoma cell, labeled with a mouse monoclonal antibody to the P . berghei circumsporozoite protein, which is associated with the surface (arrows) and with membrane-bounded vesicles (VS) in the peripheral schizont cytoplasm. N, nucleus. x 8000. (Reproduced by permission of the American Society of Tropical Medicine and Hygiene from Atkinson et al., 1989, American Journal of Tropical Medicine and Hygiene, 41, 9-17.)

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(Fig. 28). As schizonts incease in size, the density of labeling on the parasitophorous vacuole membrane decreases. Shortly before exoerythrocytic schizonts break into cytomeres between 36 and 50 h after sporozoite invasion, an unusual change in the distribution of CS protein occurs that seems to be correlated with the appearance of flocculent material in the parasitophorous vacuole space. Labeling can be detected on the limiting membranes of peripheral vesicles in the schizont cytoplasm that contain morphologically similar flocculent material (Fig. 29). The presence of CS antigen on the limiting membranes of the vesicles suggests that CS protein and flocculent material may be synthesized at the same time and exported to the surface of the parasite. Attempts to localize CS antigen in the endoplasmic reticulum and Golgi of developing schizonts with both monoclonal and polyclonal antibodies to CS protein have not been successful, however. It is possible that the vesicles may be originating from the surface of the schizont by endocytosis.

FIG.29. LR Gold section of a P . berghei exoerythrocytic schizont within a HepG2A16 hepatoma cell, labeled with a mouse monoclonal antibody to the P . berghei circumsporozoite protein. CS protein is associated with the schizont plasmalemma (arrows) and with membranes of peripheral vesicles (VS) which contain flocculent material. Label is absent from the parasitophorous vacuole membrane (PVM). The presence of label on these internal vesicles suggests that CS antigen is moving either into or out of the developing schizont. x 30 000. (Reproduced by permission of the American Society of Tropical Medicine and Hygiene from Atkinson et al., 1989, American Journal of Tropical Medicine and Hygiene, 41, 9-1 7.)

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TOXOPLASMA, SARCOCYSTIS AND EIMERIA

As with Plasmodium, pre- and post-embedding immunoelectron microscopy of coccidian parasites has focused primarily on the surface and rhoptrymicroneme antigens of merozoites and sporozoites. Sibley and Sharma (1987) reported the ultrastructural localization of T. gondii antigens which

FIG.30. Lowicryl K4M section of a Toxoplasma gondii tachyzoite, labeled with a mouse monoclonal antibody to a 55/60 kDa Toxoplasma rhoptry protein. Gold label is associated with rhoptries (R). ER, endoplasmic reticulum; N, nucleus. x 25 000. (Reproduced by permission of Elsevier Science Publishers from Sadak et al., 1988, Molecular and Biochemical Parasitology, 29, 203-21 1.) FIG.31. Epon embedded sporozoite of Eimeria tenella that had been incubated for 10 minutes with a mouse monoclonal antibody to a sporozoite surface antigen, prefixed with glutaraldehyde and then incubated with goat anti-mouse IgG colloidal gold. Colloidal gold (arrows) is associated with the surface of the sporozoite. Capping and sloughing of antibody occurred only when sporozoites were incubated with both antibody and colloidal gold before fixation. x 30000. (Reproduced by permission from Speer et al., 1985, Journal of Parasitology, 71, 3342.)

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partially protect mice from lethal challenge. Both 58 kDa and 28 kDa proteins appear to be localized primarily beneath the cell surface membrane. The distribution is somewhat different from that in Plasmodium, in which protective antigens of merozoites and sporozoites are mostly exposed on the surface of the parasites. Other studies have localized surface and rhoptrymicroneme antigens by immunoelectron microscopy (Fig. 30). Some of these antigens are secreted at the time of host-cell invasion and modify the parasitophorous vacuole membrane and space around intracellular parasites (Sibley et al., 1986; Kimata and Tanabe, 1987; Sadak et al., 1988). Yasuda et al. (1988) recently localized actin in the apical end of Toxoplasma zoites in association with microtubules, polar rings and the conoid and suggested that interaction of this cytoskeletal protein with microtubules and myosin may be important in mediating motility. Immunoelectron microscopy has been used to localize surface and refractile body antigens in species of Eimeria (Speer et al., 1983; Whitmire and Speer, 1986; Augustine and Danforth, 1987; Augustine et al., 1988). Speer et al. (1983) and Whitmire and Speer (1986) used pre-embedding immunoelectron microscopy to localize antigens on the surface of Eimeria sporozoites and on the inner walls of sporocysts and oocysts. Immunoelectron microscopy has been used in several studies to investigate the capping and shedding of immune complexes from the surface of sporozoites (Fig. 31) (Dubremetz et al., 1985; Speer et al., 1985). Only once has post-embedding immunoelectron microscopy been used to study the early stages of intracellular development of a coccidian parasite. Entzeroth et al. (1986) studied the exocytosis of a 21 kDa antigen from dense granules in the apical end of Sarcocystis muris sporozoites after host cell invasion (Fig. 32). This antigen was released into the parasitophorous vacuole space soon after sporozoite invasion, suggesting that it plays some important role in modifying this intracellular compartment. Interestingly, exocytosis did not occur through the apical pore of the rhoptry-microneme complex, but through specialized exocytotic sites in sub-apical regions of the zoite as observed in Plasmodium merozoites. C.

THEILERIA

Only a few investigators have used immunoelectron microscopy to characterize antigens and host-parasite interactions in piroplasms. Dobbelaere et al. (1985a) used a monoclonal antibody to a 68 kDa sporozoite antigen of Theileria parva to localize this molecule on the surface of sporozoites by immunoelectron microscopy. Studies of the expression of this antigen in the salivary glands of ticks demonstrated that it was synthesized by developing sporoblasts before the formation of mature sporozoites (Dobbelaere et al.,

FIG. 32. Lowicryl K4M section of a Sarcocystis muris cystozoite fixed 30 minutes after invasion of a cultured MDCK cell; section labeled with a rabbit antiserum to a 21 kDa protein. Gold label is associated with contents of dense granules (G) at the anterior end of the cystozoite. The contents of these granules are released into the parasitophorous vacuole space (arrow) around recently invaded cystozoites. x 14 300. (Reproduced by permission from Entzeroth et al., 1986, European Journal of Cell Biology 41, 182-188.) FIG. 33. Cryosection of a Theileriaparva sporozoite that had been incubated with a mouse monoclonal antibody to the sporozoite surface coat. The surface membrane (arrows) and some micronemes (M) within the sporozoite are labeled. x 45 000. (Reproduced by permission from Webster et al., 1985, European Journal of Cell Biology, 36, 157-162.) FIG. 34. Cryosection of a Theileriaparva sporozoite ( S ) . During invasion of bovine lymphocytes (L) the sporozoite surface antigen (arrow) is shed on to the lymphocyte membrane. x 45 000. (Reproduced by permission from Webster et al., 1985, European Journal of Cell Biology, 36, 157-162.)

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1985b). The antigen could be localized on nuclear membranes and cytoplasm of developing sporoblasts. As sporozoites matured, labeling was observed on the sporozoite surface and on the membranes and luminal contents of micronemes, suggesting that the micronemes may be involved in transport of this antigen to the parasite surface (Fig. 33). During invasion of bovine lymphocytes, the sporozoite surface antigen was shed and became attached to the lymphocyte surface at the site of parasite entry (Fig. 34) (Webster et al., 1985). Significantly, these workers detected the apparent exocytosis of surface antigen from micronemes, suggesting that these organelles discharge materials that may be important during host cell invasion and early intracellular development. D.

TRYPANOSOMA A N D OTHER FLAGELLATES

Trypanosomes are covered by a variant surface glycoprotein (VSG) coat which protects them against host defense mechanisms (Vickerman and Barry, 1982). The antigenic structure of the glycoprotein can be changed by a gene-switching mechanism so that the parasite is able to avoid destruction by host antibodies. Immunoelectron microscopy has been used to study the intracellular distribution and transport of VSG in blood-stream forms of T. hrucei (Duszenko et al., 1988) and T. congolense (Frevert and Reinwald, 1988) and the onset of its expression during development in tsetse flies (Glossina spp.) (Tetley et al., 1987). Intracellular transport of VSG is limited to an area between the flagellar pocket and the nucleus. VSGs are synthesized in the endoplasmic reticulum and move to Golgi complexes, tubular vesicular elements and flattened cisternae (Fig. 35). Vesicles containing VSG then move to the flagellar pocket where the protein is integrated into the surface coat (Duszenko et al., 1988). Internalization of VSG from the trypanosome surface has also been observed. Endocytotic vesicles containing VSG may form from the flagellar pocket membrane and move to fuse with lysosomes. Immunoelectron microscopy has also been used to study the intracellular distribution of tubulin in trichomonads and Giurdia (Crossley et al., 1986; Batista et al., 1988) and to characterize specific surface antigens of T. cruzi (Bretana et ul., 1986; Tachibana et al., 1986; Peyrol et al., 1987). E.

TRICHINELLA A N D OTHER NEMATODES

Most studies of nematodes in which immunoelectron microscopy has been used have employed immune sera from infected animals to identify specific antigenic targets (Prusse et al., 1983; Takahashi ez al., 1988, 1989) or to study expression of cuticular antigens or developmental changes in surface

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FIG. 35. Cryosection of a Trypanosoma brucei trypomastigote, labeled with antibody to the variant surface glycoprotein. Gold label is associated with the surface as well as portions of the Golgi apparatus, including tubulovesicular elements (arrowhead) and flattened cisternae (arrows), indicating that the trypanosome surface coat is synthesized and transported to the surface of the parasite along the classical intracellular route for glycoproteins. x 50 000. (Reproduced from Duszenko et al., 1988, Journal of Cell Biology, 106, 77-86, by copyright permission of the Rockefeller University Press.) FIG. 36. LR White section of a Trichinella spiralis muscle larva, incubated with rat immune serum to T . spiralis. Gold label is associated with possible protective antigens in the inner layers of the cuticle (C). x 17 000. (Reproduced by permission from Takahashi et al., 1988, Journal of Parasitology, 74, 27C274.)

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antigens of oocytes and spermatozoa (Bradley and Burghardt, 1976; Wu and Foor, 1980; Kiefer et al., 1986). Takahashi et al. (1988a,b) used immune serum from rats that had been infected with Trichinella spirulis to identify organelles that may be involved in the development of protective immunity (Fig. 36). Immune targets were associated with the surface and hypodermis of the cuticle, material within the esophagus and midgut, antigenic components of the hemolymph, stichocyte granules, and aggregates of glycogen in a variety of cells. The authors concluded that only a limited number of parasite constituents are antigenic in rats. Most of these are associated with surfaces of the parasites that are exposed to the immune system or to materials that may be released from intact worms as excretory/secretory products. F.

SCHISTOSOMA AND FASCIOLA

Trematode infections are a major cause of morbidity and mortality in humans and domestic animals. Recent work on Schistosoma and Fasciola has focused on the characterization of tegumental and excretory/secretory antigens that may have protective or diagnostic value. Several studies have localized protective antigens in the tegument of S. mansoni (Matsumoto et al., 1988b; Taylor et al., 1988). A 28 kDa antigen which has glutathione transferase activity has been identified in excretory epithelial cells, tegumental and sub-tegumentary parenchymal cells, and tegumental granules in the head glands of adult worms (Taylor et al., 1988). Matsumoto et al. (1988b) localized paramyosin and actin in the tegument of S. mansoni by immunoelectron microscopy (Figs 37 and 38). Actin was localized in the paracrystalline cores of tegumental spines (Fig. 37) and in cortical muscle beneath the tegument (Matsumoto et al., 1988b). By contrast, paramyosin was restricted to the tegument where it was unexpectedly found in membrane-bound elongate bodies (Fig. 38). The presence of this cytoskeletal protein in the FIG.37. LR Gold section of an adult Schistosoma mansoni, labeled with an antibody to chicken gizzard actin which recognizes all actin isoforms. This cytoskeletal protein is a major constituent of the tegumental spines (TS) and is also present in cortical muscle (arrow) beneath the tegument. x 30 000. (Reproduced by permission from Matsumoto et af., 1988, Nature, 333, 7 6 7 8 . Copyright 0 1988 Macmillan Magazines Ltd) FIG.38. LR Gold section of an adult Schistosoma mansoni, labeled with a rabbit antiserum to paramyosin. This invertebrate cytoskeletal protein is associated with non-filamentous, membrane-bound elongate bodies within the tegument (T), cytons and the cytoplasmic tubes (arrow) which connect the tegument to cytons. x 15 180. (Reproduced by permission from Matsumoto et af., 1988, Nature, 333, 7 6 7 8 . Copyright 0 1988 Macmillan Magazines Ltd) Inset: gold particles associated with elongate bodies of the tegument. x 54 000.

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outermost layer of the schistosome body may explain why vaccination with this protein induces protective immunity in mice (Lanar et al., 1986). Other tegumental antigens have been localized by immunoelectron microscopy within secretory granules, Hanna et al. (1988) identified glycocalyx antigens in tegumental granules of F. hepatica. These granules are important in renewal of the glycocalyx after it is shed in the presence of immune serum. Sloughing of the tegumental glycocalyx appears to be an important mechanism for evading the host immune response. Studies of excretory/secretory antigens have localized these molecules in the contents and epithelium of the intestinal tract (Fujino et al., 1985; De Water et al., 1986a,b), in the excretory system (i.e. flame cells) (De Water et al., 1987a), and in the gland cells and epidermis of miracidia (Bogitsh and Carter, 1975). De Water et al. (1987b) used immunoelectron microscopy to study the fate of circulating anodic and cathodic antigens in the livers of mice infected with S. mansoni. These workers found schistosome antigen and immune complexes within secondary lysosomes in Kupffer cells, granuloma macrophages, and endothelial cells from infected livers. Thus, the technique may prove to be important for studying immunopathological processes in infected hosts. Immunoelectron microscopy has also been used to study the acquisition and mimicry of host antigens by trematodes (McLaren and Smithers, 1975; Yoshino and Cheng, 1978). McLaren and Smithers (1975) used an immunoperoxidase technique to demonstrate that young schistosomes acquire host antigens within several hours after host penetration. Yoshino and Cheng (1978) used a similar procedure to show that surface antigens of miracidia may mimic hemolymph proteins of their snail hosts. IV. CONCLUSIONS During the past decade, techniques of immunology and molecular biology have been used to identify and isolate specific parasite proteins that may be capable of producing protective immunity against protozoan and helminth parasites. Immunoelectron microscopy is a powerful tool for studying hostparasite interactions and it is playing an increasingly important role in identifying specific immune targets and characterizing the precise subcellular localization, transport, and expression of parasite antigens. In addition, this technique can help to clarify specific functions of subcellular organelles, which may not otherwise be detected by standard electron microscopy or biochemical techniques. In studies of Plasmodium, immunoelectron microscopy has been especially valuable in characterizing the antigenic composition of intracellular compartments, e.g. parasitophorous vacuole and

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cytoplasmic clefts, that cannot be isolated, purified, and studied by current biochemical procedures. Therefore, immunoelectron microscopy will undoubtedly contribute to a better understanding of the relationship between structure and function in parasites. Technical aspects of immunoelectron microscopy have improved significantly in recent years, making these procedures practicable for most electron microscopy facilities. At the same time, important new advances in the field have not yet been applied to the study of parasites. The development of cryofixation and freeze-substitution techniques (Linner et al., 1986) and the application of immunocytochemistry to freeze-fracture techniques (Forsman and Da Silva, 1988) show considerable promise for studying molecular and ultrastructural interactions between hosts and parasites. As technical barriers to performing immunoelectron microscopy diminish, these techniques will have many applications in the field of parasitology. ACKNOWLEDGEMENTS This work was supported in part by grants from the US Agency for International Development (DPE-O453-A-00-4027), USPHS (AI-10645), UNDP/World Bank/WHO Special Programme for Research and Training in Tropical Diseases and US Army R & D Command Contract no. DAMD17-85-C-5179. This is contribution no. 1852 to the Army Research program on antiparasitic drugs. The authors thank Drs G. A. M. Cross, J. F. Dubremetz, Y. Matsumoto, D. Sibley, C. Speer, Y. Takahashi, M. Torii, S. Uni and P. Webster for providing electron micrographs for this chapter. Also, we acknowledge the technical assistance of Kiet Dan Luc and Lucy Sanders. REFERENCES

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Sibley, L. D., Krahenbuhl, J. L., Adams, G. M. W. and Weidner, E. (1986). Toxoplasma modifies macrophage phagosomes by secretion of a vesicular network rich in surface proteins. Journal of Cell Biology 103, 867-874. Siddiqui, W. A., Tam, L. Q., Kramer, K. J., Hui, G. S. N., Case, S. E., Yamaga, K. M., Chang, S. P., Chan, E. B. T. and Kan, S.-C. (1987). Merozoite surface coat precursor protein completely protects Aotus monkeys against Plasmodium falciparum malaria. Proceedings of the National Academy of Sciences of the USA 84, 3014-301 8. Singer, S. J. (1959). Preparation of an electron dense antibody conjugate. Nature 183, 1523-1 524.

Slot, J. W. and Geuze, H. J. (1981). Sizing of protein A-colloidal gold probes for immunoelectron microscopy. Journal of Cell Biology 90, 533-536. Slot, J. W. and Geuze, H. J. (1984). Gold markers for single and double immunolabelling of ultrathin cryosections. In “ Immunolabelling for Electron Microscopy” (J. M. Polak and I. M. Varndell, eds). Pp. 129-142. Elsevier Science Publishers, New York. Slot, J. W. and Geuze, H. J. (1985). A new method of preparing gold probes for multiple-labeling cytochemistry. European Journal of Cell Biology 38, 87-93. Smythe, J. A., Coppel, R. L., Brown, G. V., Ramasamy, R., Kemp, D. J. and Anders, R. F. (1988). Identification of two integral membrane proteins of Plasmodium falciparum. Proceedings of the National Academy of Sciences of the USA 85, 5195-5199. Somogyi, P. and Takagi, H. (1982). A note on the use of picric acid-paraformaldehyde-glutaraldehyde fixative for correlated light and electron microscopic immunocytochemistry. Neuroscience 7 , 1779-1 783. Speer, C. A., Wong, R. B. and Schenkel, R. H. (1983). Ultrastructural localization of monoclonal IgG antibodies for antigenic sites of Eimeria tenella oocysts, sporocysts and sporozoites. Journal of Protozoology 30,548-554.

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TECHNICAL APPENDIX Immunoelectron microscopy exposes fresh tissues to fixatives, buffers, organic solvents and embedding media that may extract molecules and cause denaturation and loss of antigenicity. For abundant antigens or antigens concentrated in discrete compartments, large losses of immunoreactivity during processing may still allow good ultrastructural localization. By contrast, labile or sparsely distributed antigens may be difficult to localize with any of the methods that are commonly used. We use procedures developed by Tokuyasu (1 986) for preparing and labeling cryosections, but have developed our own modifications for embedding, sectioning and labeling when using LR White and LR Gold resins. A.

FIXATION

We have experimented with a variety of different fixatives, but have had best results with a mixture of 1YO paraformaldehyde and 0.2% glutaraldehyde in 0.1 M phosphate buffer, pH 7.4. This fixative preserves relatively good ultrastructure and preserves the antigenicity of a wide variety of molecules. Prepare as follows: (i) Mix 1.O g of paraformaldehyde powder with 45 ml of double-distilled water, cover with aluminum foil, and heat with stirring on a hot-plate at 60°C for 2Ck 30 min. (ii) Add 4-6 drops of 1 N NaOH. The milky white solution should clear instantly. (iii) Cool to room temperature and mix with 50 ml of 0.2 M phosphate buffer, pH 7.4.

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(iv) Add 0.8 ml of the 25% stock glutaraldehyde solution and mix. (v) Check pH, bring total volume to 100 ml, and cool. The fixation time and temperature will depend on the tissue. We routinely fix Plasmodium-infected erythrocytes for 10 min at room temperature. B.

EMBEDDING AND SECTIONING WITH LR WHITE AND LR GOLD RESINS

LR White resin is a premixed acrylic formulation which is polymerized by heat or by addition of an initiator for a rapid “cold” cure (Newman and Hobot, 1987). LR White was originally designed for use at room temperature, but we have found that it will infiltrate tissues at temperatures as low as -20°C. At temperatures between - 20°C and - 30°C the resin will solidify. LR White is used to infiltrate tissues at low temperatures to protect antigens from the denaturing effects of ethanol (Armbruster el al., 1983) and the resin is then polymerized at 40°C for 5 days. This temperature is lower than that recommended by the manufacturer, but still adequate to produce blocks that are easy to section. Undercuring the resin at lower temperatures seems to increase the immunoreactivity of the embedded sample, perhaps by reducing intermolecular cross-linking and allowing better penetration of immunoreagents. LR Gold resin was designed for use at temperatures as low as -25°C and we normally use this resin for tissues that are not darkly pigmented or rich in hemoglobin. LR Gold resin preserves antigenicity of some molecules better than Lowicryl, but may not preserve ultrastructure as well (Yokota and Oda, 1985). We have had only limited success in producing well-polymerized blocks of erythrocytes with resin that are polymerized with ultraviolet light, e.g. Lowicryl and LR Gold. The high oxygen content and dark color of hemoglobin-rich erythrocytes may inhibit polymerization of the resin or limit penetration of ultraviolet light. Sectionable blocks can be produced, however, if erythrocytes are polymerized as a suspension rather than as a pellet. LR Gold can be polymerized at low temperatures after addition of appropriate initiators (benzoin methyl ether, benzoil or camphoquinone) and exposure to intense ultraviolet or blue light. We routinely use benzoin methyl ether as an initiator at concentrations of O . M . 7 5 % (w/v) and polymerize the tissue at -20°C under intense ultraviolet light for 24-48 h. Higher initiator concentrations, i.e. 0.75%, require shorter polymerization times and seem to produce blocks with better sectioning qualities. Tissues should be dehydrated as rapidly as possible to minimize exposure to alcohols. Actual times will depend on the size and density of the tissue. When tissue is embedded in LR Gold, initiator is dissolved in the resin at room temperature and the mixture is then chilled at -20°C. Ethanol solutions and resin mixtures should be made beforehand and cooled to -20°C before use. (i) 30% Ethanol; 10-20 min; 4°C. (ii) 50% Ethanol; 10-20 min; -20°C. (iii) 70% Ethanol; 10-20 min; -20°C. (iv) 1 part 95% Ethanol, 2 parts resin; 10-20 min; -20°C.

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(v) 100% resin; 60min; -20°C. (vi) 100% resin; 60 min; -20°C. (vii) 100% resin; 60min; -20°C. (viii) Fresh resin overnight. For LR White embedding, samples are left overnight at 4°C. For LR Gold embedding, samples are left overnight at -20°C. (ix) Transfer samples to gelatin capsules and cap tightly to exclude as much oxygen as possible. LR White is polymerized at 40°C for five days. LR Gold is polymerized for from 24 h to several days at -20°C under ultraviolet light. Polyethylene Beem capsules can be used to hold the tissue, but the surface of the blocks may be tacky after polymerization. Polymerized blocks of LR White and LR Gold resin are brittle, but may be sectioned easily with glass or diamond knives. Sections are picked up on unsupported 200-400 mesh nickel grids. Formvar-coated grids may be used, but sections tend to wrinkle during immunostaining. Since subsequent immunolabeling steps use solutions which contain detergents (Tween 20), sections often detach and float away unless they are bound firmly to the grids. We make grids “sticky” by coating the metal surfaces of the grids with formvar before they are used to pick up sections. We prepare the grids by dipping them into a 0.5% (w/v) solution of formvar in dichloroethane and then rapidly draining away all excess solution on a piece of highly absorbent filter paper. It is important that the grids be placed flat on the filter paper before the formvar solution dries; otherwise, thick deposits of plastic will obscure the spaces between grid bars.

C.

I.

LABELING PROCEDURES FOR LR WHITE AND LR GOLD SECTIONS

Blocking

Fixation with glutaraldehyde can cause the non-specific binding of ligands and ligand-marker conjugates to unquenched aldehyde groups in tissue sections (Hodges ct al., 1984). Most authors suggest that free aldehyde groups should be blocked before immunolabeling with solutions that contain free amino groups to reduce nonspecific background binding. This step is especially important in pre-embedding immunocytochemistry and cryo-ultramicrotomy where tissues or tissue sections are incubated with antibodies and electron-dense markers before embedding. Suitable blocking agents include buffered solutions of gelatin, serum albumin, non-fat dried milk, 0.1 M glycine, 0.1 M lysine, 0.05 M ammonium chloride or 0.05 M ammonium carbonate, and solutions of 0.05-1 .O% sodium borohydride (Lillie and Pizzolato, 1972; Weber et al., 1978; Farr and Nakane, 1981; Van Leeuwen, 1982; Eldred et al., 1983). Some authors have suggested that treatment with sodium borohydride before embedding may help to restore antigenicity to glutaraldehyde-fixed tissue (Eldred et al., 1983). We use solutions of 5% non-fat dried milk to block thin sections of resinembedded material.

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2.

213

Etching

Etching techniques unmask antigenic sites that may have been destroyed or altered during dehydration and embedding. Strong oxidizing agents such as sodium metaperiodate and hydrogen peroxide are often used on epoxy-embedded tissues before immunolabeling to increase the hydrophilic nature of the sections, unmask antigenic sites and improve their reactivity with aqueous solutions of antibodies (Bendayan and Zollinger, 1983). This treatment oxidizes hydrophobic alkane side chains in the resin to alcohols, aldehydes and acids, thereby making the resin surface more hydrophilic (Causton, 1984). Since this treatment may also oxidize tissue-associated antigens and affect their reactivity with antibodies, it should be used with caution. Most of the acrylic resins now in use, e.g. LR White, LR Gold and Lowicryl, are hydrophilic and do not require this treatment. We have found, however, that treatment of LR White or LR Gold sections with a saturated aqueous solution of sodium metaperiodate may improve the immunoreactivity of some resin-embedded antigens. Actual removal of resin from grid-mounted sections with solutions of sodium ethoxide may significantly enhance immunolabeling (Mar and Wight, 1988). lngram et al. (1988) have successfully used this technique in combination with sodium metaperiodate to unmask antigens in erythrocytes embedded in LR White resin.

3. Labeling A variety of techniques for labeling grid-mounted resin sections has been described. Many workers use individual drops of reagents on wax or parafilm sheets, porcelain plates with 6-12 individual wells, or multi-well microtiter plates for holding washing and incubation solutions. We have found that the Hiraoka staining kit (Polysciences, Inc.) offer a number of advantages over other procedures. This device consists of a flexible plate of nalgene with precisely cut rows of shallow slits. Grids are firmly mounted in a vertical position on the plate by their rims, where they can be incubated in drops of blocking solutions, antibodies and colloidal gold and “jet-washed” gently with a Pasteur pipette or dipped into beakers that contain appropriate washing solutions. This procedure allows both sides of grid-mounted resin sections to be labeled simultaneously, to increase the density of immunolabeling. In addition, the plates are much easier and faster to handle than individual grids and allow large numbers of grids to be incubated and washed at the same time. The procedure is described below. (i) Erch grids in drops of saturated aqueous sodium metaperiodate for 3&60 min and rinse with distilled water. This step is optional and may destroy the immunoreactivity of some antigens. (ii) Cover grids with drops of PBS-milk-Tween blocking solution ( 5 % non-fat dried milk in 0.1 M phosphate buffer, pH 7.4, with 0.01 YOTween 20 and 0.9% NaCI) and incubate for 30 min.

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(iii) Remove blocking solution and cover grids with drops of the primary antibody diluted with PBS-BSA-Tween (IYo bovine serum albumin, fraction V, in 0.1 M phosphate buffer, pH 7.4, with 0.01% Tween 20 and 0.9% NaCI). The primary antibody can also be diluted in the PBSmilk-Tween blocking solution to control problems with non-specific background labeling. While this works well with some antibodies, it may completely abolish labeling with others. Optimal antibody dilutions must be determined by trial and error, although we have had good results with antibody concentrations ranging from 2 to 200 pg ml. - Incubate for 2 h at room temperature or overnight at 4°C in a humidity chamber. (iv) Jet-wash grids with PBS-BSA-Tween and incubate with three 5-min changes of PBS-BSA-Tween. (v) Incubate grids with rabbit anti-mouse IgG secondary antibody diluted to approximately 2 M O pg ml- with PBS-BSA-Tween. The optimal dilution must be found by trial and error, but should remain the same for each lot of antibody. Incubate the grids for 1 h at room temperature in a humidity chamber. (vi) Jet-wash grids with PBS-BSA-Tween and incubate with three 5-min changes of PBS-BSA-Tween. (vii) Incubate grids with protein A-gold or goat anti-rabbit antibody-gold diluted 1/20 with PBS-BSA-Tween for 1 h at room temperature in a humidity chamber. To control background problems, immunoglobulin-gold can also be diluted in the PBSmilk-Tween blocking solution. (viii) Jet-wash grids with 0.1 M phosphate buffer, pH 7.4, and fix grids in drops of 2.5% glutaraldehyde in 0.1 M phosphate buffer, pH 7.4, for 15 min to stabilize the bound gold. (ix) Jet-wash with double-distilled water and dry with filter paper. (x) Stain the grids for 30min in 2 % (w/v) uranyl acetate dissolved in 50% methanol. Jet-wash the grids with 50% methanol and dry. (xi) Stain sections with Reynold's lead citrate for 5 min, wash with double-distilled water and dry. (xii) Coat grids lightly with carbon in a vacuum evaporator to stabilize the sections.

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